Strains

- Wild type isolates tested for RNAi: Caenorhabditis elegans N2, CB4856, JU1580, C. briggsae AF16, JU1264 and HK104 (data not shown, Amélie Broucke), C. remanei PB4641, C. brenneri PB2801, C. drosophilae DF5077, C. plicata SB355, C. sp. 2 DF5070, C. angaria RGD1, C. sp. 5 JU727, C. sp. 6 EG4788 (gift of M. Ailion), C. sp. 7 JU1199 and JU1593, C. sp. 8 QX1182 (gift of M. Rockman), C. sp. 9 JU1325, C. sp. 10 JU1333, C. sp. 11 JU1373 (V. Robert and M.-A. Félix), C. sp. 12 JU1427 (P. Châtelet and M.-A. Félix), C. sp. 13 JU1528, C. sp. 14 EG5716 (gift of M. Ailion), C. sp. 15 QG122 (gift of M. Rockman), C. sp. 16 JU1873 (J.-B. Pénigault), C. sp. 17 JU1825 (C. Braendle and M.-A. Félix), C. sp. 18 JU1857 (C. Braendle and M.-A. Félix), C. sp. 19 EG6142 (gift of M. Ailion), C. sp. 20 NIC113 (gift of C. Braendle)

- Additional wild type isolates tested for transformation: C. remanei SB146, C. brenneri CB5161, C. angaria PS1010; C. plicata SB355, C. sp. 1 SB341 (gift of W. Sudhaus).

- Transgenic strains (see construction below): C. briggsae JU1018 (mfIs42[Cel-sid-2; Cel-myo-2::DsRed2]); JU1076 (mfIs48[Cel-sid-2; Cel-myo-2::DsRed2]); C. remanei JU1184 (mfEx34[Cel-sid-2; Cel-myo-2::DsRed2], likely integrated); C. sp. 9 JU1591 (mfEx65[Cel-sid-2; Cel-myo-2::DsRed2]); C. sp. 11 JU1592 (mfEx66[Cel-sid-2; Cel-myo-2::DsRed2]) and JU1874 (mfEx44[Cel-sid-2; Cel-myo-2::GFP]).

Generation of dsRNA-expressing bacteria

- Oti-actin RNAi plasmid: An Oti-actin fragment was isolated using primers designed using the C. elegans actin sequence: primers actinI (ATGTGCAAGGCCGGATTCG) and actinIII (ATAGGGACGGTGTGG). The PCR was performed on genomic DNA of the Oscheius tipulae CEW1 strain with an annealing temperature of 55°C and 2.5 mM MgCl₂ [18]. The amplified fragment was inserted into the pGEMTeasy plasmid, yielding pMD13. After digestion of pMD13 by NotI, the Oti-actin fragment was inserted into pPD129.36 [3] and the resulting pMA33 plasmid transformed into chemically competent HT115 cells used for RNAi experiments. The sequence of the Oti-actin fragment amplified with actinI and actinIII is:

“ATGTGCAAGGCCGGATTCGCCGGAGACGATGCTCCTCGCGCCGTCTTCCCCTCCATCGTCGGTCGCCCCCGTCACCAGGGAGTCATGGTCGGAATGGGACAGAAGGACAGCTACGTCGGAGACGAGGCCCAGTCCAAGAGAGGTATCCTGACCCTCAAGTACCCCATTGAGCACGGTATCGTCACCAACTGGGATGACATGGAGAAGATCTGGCACCACACCTTCTACAATGAGCTCCGTGTCGCCCCTGAGGAGCACCCCGTCCTCCTGACTGAGGCTCCCCTCAATCCCAAGGCTAACCGTGAGAAGATGACCCAGATCATGTTCGAGACCTTCAACGCTCCCGCCATGTACGTCGCCATCCAGGCCGTCCTCTCCCTCTACGCCTCCGGACGTACCACCGAGTCGTCCTCGACTCTGGAGATGGAGTTACCCACACCGTCCCTAT”.

The matching actin gene sequences of C. elegans, C. briggsae, C. remanei, C. brenneri and C. angaria are available in Wormbase (www.wormbase.org). Those of the other species have not been determined.

- Cre-unc-22 RNAi plasmid: A Cre-unc-22 fragment was amplified using primers oMA197 (AGGCGGCCGCGTCGAAGAAGACGACCAAGC) containing a NotI restriction site and oMA198 (TGCCATGGTCCAGCTTCGCTTCAATTTT) containing a NcoI restriction site. This PCR fragment was digested with NcoI and NotI, and inserted into pPD129.36 plasmid, previously digested with NcoI and NotI. Genomic sequences of Caenorhabditis species can be found in Wormbase (www.wormbase.org).

- Can-ama-1 and Cdr-ama-1 RNAi plasmids: A Can-ama-1 fragment was amplified from C. angaria genomic DNA using primers oIN114 (GGGGACAAGTTTGTACAAAAAAGCAGGCTCCCTACACTGAAGATGTGATC) and oIN115 (GGGGACCACTTTGTACAAGAAAGCTGGGTGTGTTGTATGCTCCAATTTGC). A Cdr-ama-1 fragment was amplified from C. drosophilae genomic DNA using primers oIN124 (GGGGACAAGTTTGTACAAAAAAGCAGGCTGCGATGGGAGGTCGTGAAG) and oIN125 (GGGGACCACTTTGTACAAGAAAGCTGGGTCGAATCACATCTTCCGAG). oIN114 and oIN124 contain an attB1 forward site, while oIN115 and oIN125 contain an attB2 reverse site. Fragments oIN124-oIN125 and oIN114-oIN115 were inserted into pDONR221 according to BP reaction's protocol (Multisite Gateway Invitrogen) and a LR reaction was performed to insert the ama-1 fragment into the pDEST L4440 Gateway feeding vector. Chemically competent HT115 cells were transformed with these Gateway RNAi ama-1 feeding plasmids. ama-1 codes for RNA polymerase II and is called pol-2 in [5]. We use here the Wormbase name.

- Cbr-lin-12 RNAi plasmid: A fragment of Cbr-lin-12 gene was amplified with oMA158 (AGccatggTACTTGTTCTTC) and oMA159 (TGgcggccgcTTCTCACTGAACATT). This PCR fragment was digested with NcoI and NotI, and inserted into the pPD129.36 plasmid previously digested with NcoI and NotI, yielding pMA52.

- Cel-rol-6, Cel-pos-1 and Cel-unc-22 RNAi clones were from the Ahringer library [19].

dsRNA feeding protocol

Bacteria were grown overnight in LB with 20 µg/mL ampicillin and 12.5 µg/mL tetracyclin. They were then seeded onto 55 mm diameter NGM agar plates containing 1 mM IPTG, 20 µg/mL ampicillin and 12.5 µg/mL tetracyclin. Seeded plates were incubated at room temperature for 2 days.

For hermaphroditic species, two RNAi plates with three L4-stage hermaphrodites were incubated at 23°C for 24 hrs. Then, for each plate, each animal was isolated onto a new RNAi plate seeded with the same bacteria, allowed to lay eggs overnight (ca. 16 hrs) at 23°C and then removed. Its progeny was scored after 24 hrs at 23°C for phenotypes related to actin RNAi and 48 hrs for phenotypes related to unc-22 RNAi.

For male-female species, eight L4-stage females and four males were incubated at 23°C for 24 hrs on NGM plates seeded with HT115 bacteria expressing dsRNAs for the gene of interest. Then, six fertilized females were isolated onto a new RNAi plate and allowed to lay eggs for 16 hrs at 23°C before being removed.

For Oti-actin RNAi experiments, the progeny was scored two days later with a stereomicroscope for number of wild-type larvae, number of deformed larvae (L1s and later stages), and number of dead embryos. For Cel-pos-1 and ama-1, the progeny was scored for number of wild-type larvae and number of dead embryos. For unc-22, we scored three days later the twitching phenotype of the adult progeny.

Generation of dsRNAs for injections

PCR products corresponding to a part of the Oti-actin or Cel-unc-22 genes were generated with T7 primer 5′-AATACGACTCACTATAG-3′ [20] from the Oti-actin or Cel-unc-22 RNAi clones, using the following cycle conditions: 95°C 50 sec, 52°C 30 sec, 72°C 90 sec for 25 cycles. Purification of the PCR product was performed with Nucleospin Extract II kit (Macherey Nagel). Oti-actin or Cel-unc-22 dsRNAs were then generated using the MEGAscript kit (Ambion).

dsRNAs were injected at 50 ng/µL into both gonadal arms of wild-type young adult hermaphrodites or females - depending on the reproductive mode of the species. For hermaphroditic species, the injected animal was transferred to a fresh plate 24 hrs after injection, and cultured at 23°C. For male-female species, a male was added onto the plate with the injected female and transferred with the female the day after. Phenotypes were scored as described above.

Generation of Cel-sid-2 transgenic animals

A Cel-sid-2 PCR product was obtained from N2 genomic DNA using primers oMA183 (GCTCAAAACCAACCTTAACTGC) and oMA184 (TCTTGCATGGTCCCCAAGTA). This product was injected into C. briggsae AF16, C. remanei PB4641 and C. sp. 11 JU1373 at a concentration of 15 ng/µL, mixed with pWD47 (Cel-myo-2::DsRed2) at 10 ng/µL and carrier pBS to a final concentration of 150 ng/µL.

C. briggsae JU1018 and JU1076 are integrated lines produced by γ-irradiation of the transgenic strain JU977 (mfEx32[Cel-sid-2; Cel-myo-2::DsRed2]), backcrossed five times to AF16.

Transgenesis in different Caenorhabditis species

The syncytial female germ line of different Caenorhabditis species was injected using the same protocol as in C. elegans [21], [22], injecting both gonadal arms when possible. Each injected animal was then transferred singly to a culture plate (except in the C. plicata experiment where two females were placed on each plate to reduce plate consumption). For male-female species, males were added to the plate with the injected female. The marker phenotype was assayed in the F1 and F2 progeny of each injected parent. The proportion of injected parents that show positive F1 or F2 progeny, respectively, is indicated in Table 1.

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Table 1. Transgenesis of Caenorhabditis species.

https://doi.org/10.1371/journal.pone.0029811.t001

The pJL53 plasmid (Cel-hsp-16-18::GFP) is a gift of Jean-Louis Bessereau (see [23] for the corresponding cis-regulatory sequences). To assay its expression, animals were heat-shocked at 33–37°C for 1.5–2 hrs and allowed to recover at 20°C for 1–2 hrs before scoring GFP fluorescence. The pRF4 plasmid contains a gain-of-function version of the rol-6 collagen gene, rol-6(su1006), which causes a dominant Roller phenotype [21]. The pTG96 (Cel-sur-5::GFP) plasmid is a gift from Min Han's laboratory and is expressed in most somatic cells [24]. The pPD118.33 (Cel-myo-2::GFP) plasmid is a gift from Andy Fire's laboratory and is expressed in the pharynx [25], [26]. The Cel-myo-2::DsRed2 pWD47 plasmid is a gift of Wayne Davis and Erik Jorgensen.

Statistical analyses

All statistical tests were performed with R (http://www.r-project.org/). A Mann-Whitney-Wilcoxon test was used to compare the numbers of wild-type larvae in the actin and ama-1 RNAi experiments. An exact Fisher test was used on Cre-unc-22 RNAi experimental results.

Devising an RNAi test using a highly conserved gene

RNA interference requires a high degree of sequence similarity between the administered dsRNAs and the target gene. We looked for a dsRNA sequence that we could use in all Caenorhabditis species, and beyond the genus. Actin was a good candidate, being a highly conserved, abundant and essential cytoskeletal protein. Indeed, the actin genes are highly conserved at the nucleotide level, presumably because of their high level of expression, resulting in a strong codon usage bias, i.e. the nucleotides at synonymous positions tend to be conserved among closely related species and gene copies. In order to reduce a possible bias in efficiency of RNAi among Caenorhabditis species, we chose to score their respective sensitivity using an actin gene of a species outside the genus, namely Oscheius tipulae CEW1 [27] (see [28] for its phylogenetic position). We cloned an Oti-actin gene fragment in a plasmid containing two inducible T7 promoters (pPD129.36) that flank the gene fragment in mirror orientation.

Inactivation of actin gene expression in C. elegans [29] results in a spectrum of phenotypes that appear during the course of the experiments, as a function of time and severity of defect. The mildest effect concerns deformation of the body of juveniles and adults grown on bacteria expressing Oti-actin dsRNAs (Figure 1), followed in order of increasing severity by a delayed growth of the juveniles, an embryonic arrest and finally sterility of the mother as the strongest and latest phenotype.

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Figure 1. Outline of the RNAi test.

(A) Experimental design of the RNAi test. a) L4 larvae were transferred on day 1 onto two RNAi plates seeded with bacteria producing Oti-actin dsRNAs. b) On day 2 (24 hrs later), the animals were isolated onto a new RNAi plate seeded with the same bacteria. c) On day 3 (after 16 hrs), the adults were removed. d) On day 4, the progeny was scored for the number of wild-type larvae, deformed larvae and embryos (these embryos must be highly delayed, arrested or dead). Experiments were performed at 23°C. (B) Morphological phenotype in the actin RNAi experiments. a) Deformed L1 larva of the C. elegans N2 strain on HT115 bacteria expressing Oti-actin dsRNAs (visualized here using Nomarski microscopy but visible under a dissecting microscope). b) Control L1 larva of the C. elegans N2 strain grown on control HT115 bacteria expressing Cbr-lin-12 dsRNAs.

https://doi.org/10.1371/journal.pone.0029811.g001

We devised a test of Oti-actin RNAi efficiency using these different phenotypic effects (Figure 1). We first incubated young adult animals for one day on dsRNA-expressing bacteria, transferred them to a new plate, allowed them to lay for a further 16 hours at 23°C and removed them. We then scored the progeny over the next days for the following traits: number of arrested embryos, number of deformed larvae, number of normal larvae. Most importantly, the number of progeny represents the fertility level of the mother. As a negative control, we used a clone of bacteria expressing dsRNAs that were not recognized by the tested species (e.g. Cel-rol-6 or Cbr-lin-12 when appropriate). In addition, C. elegans N2 was used as an internal positive control for each experiment.

Evolutionary variation in sensitivity to external Oti-actin dsRNAs in the Caenorhabditis genus

We systematically assayed all species of Caenorhabditis in culture for their sensitivity to external application of dsRNA (except for C. japonica strains, which have a very poor fecundity [5]; our observations). We first confirmed in our assay previous results that had been obtained by soaking the animals in dsRNAs corresponding to the species ama-1/pol-2 (coding for the RNA polymerase II large subunit) gene [5]. We further scored many new species, some of which were found to be naturally sensitive to external dsRNAs. Figure 2 shows examples of a species that is insensitive to ingested Oti-actin dsRNAs, Caenorhabditis sp. 18 (Figure 2A), and of a highly sensitive species, Caenorhabditis sp. 15 (Figure 2B). Further results are reported in Table S1 and Figure 3.

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Figure 2. Comparison of sensitivity to ingested Oti-actin dsRNAs of C. elegans N2 to two examplary species.

(A) C. sp. 18 JU1857 is insensitive to RNAi by feeding, whereas C. elegans N2 shows a response. C. sp. 18 JU1857 and C. elegans N2 progeny were scored using the experimental design described in Figure 1, using Oti-actin and Cbr-lin-12 (negative control). (B) C. sp. 15 QG122 is highly sensitive to ingested Oti-actin dsRNAs. Here, feeding of Oti-actin dsRNA resulted in complete sterility of the mothers, whereas in the same experiment, treated N2 mothers produced arrested embryos and larvae. Statistical comparisons were made to the results of control experiments with Cbr-lin-12 dsRNA, which has no effect on either strain, using a Mann-Whitney-Wilcoxon rank sum test on the number of normal larval progeny of each parent. The significance of the difference is depicted as follows: (NS) non-significant, (*) 0.01<p<0.05, (**) 0.001<p<0.01, (***) p<0.001. Error bars indicate the standard error of the mean over individuals (n = 6–12, see Table S1 for details).

https://doi.org/10.1371/journal.pone.0029811.g002

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Figure 3. Evolution of RNAi sensitivity in the Caenorhabditis genus.

The RNAi results are displayed on the phylogeny from [17], for the different dsRNA administration methods and for different genes. The red stars indicate a loss of sensitivity to ingested Oti-actin dsRNAs in a parsimonious evolutionary scenario (see text). Note that the absence of an external Oti-actin RNAi response may correspond to an inability to respond either to all ingested dsRNAs (as in C. briggsae and likely in C. drosophilae), or only to specific dsRNAs (as in C. elegans JU1580 and C. angaria). In blue: results from [13]. In green: result`s from [5]. The dsRNAs for the ama-1 gene were specific to the tested species. We did not find differences among different tested isolates of C. briggsae and C. sp. 7 (see Materials and Methods for the identity of other tested strains), nor did Winston et al. for different isolates of C. brenneri [5]. C. brenneri can respond to ama-1 dsRNAs if they are injected into the gonad, indicating that this species is defective in the systemic spread of the signal from body cavity to gonad. I: Elegans supergroup of Caenorhabditis species; II: Drosophilae supergroup as in [17].

https://doi.org/10.1371/journal.pone.0029811.g003

In addition to C. elegans and C. sp. 1 SB341, we found that Caenorhabditis sp. 6 EG4788, C. sp. 7 JU1199, C. sp. 13 JU1528, C. sp. 14 EG5716 and C. sp. 15 QG122 were naturally sensitive to ingested dsRNAs and thus easily amenable to RNAi screens. C. sp. 16 JU1873, C. sp. 20 NIC113 and perhaps C. plicata appeared mildly sensitive to ingested Oti-actin dsRNAs. In contrast, C. briggsae, C. remanei, C. brenneri, C. drosophilae, C. sp. 2 DF5070, C. sp. 3 RGD1, C. sp. 5 JU727, C. sp. 8 QX1182, C. sp. 9 JU1325, C. sp. 10 JU1333, C. sp. 11 JU1373, C. sp. 12 JU1428, C. sp. 17 JU1825, C. sp. 18 JU1857 and C. sp. 19 EG6142 were insensitive. Insensitivity to ingested Oti-actin dsRNAs is thus predominant in terms of number of species. Importantly, it does not follow a phylogenetic distribution (see below) that would suggest that it can be explained by the sequence divergence of the actin gene of the species relative to the Oti-actin gene fragment.

It is noteworthy that C. elegans N2 RNAi efficiency is not the strongest in this assay. Especially, milder phenotypes than full mother sterility, such as deformed larvae, embryonic lethality or retarded development, were often observed using C. elegans N2; this is also the case with C. sp. 20 but not other sensitive species where the effect of actin RNAi was stronger (Figure 2 and Table S1). C. elegans N2 RNAi efficiency is thus in a range that appears highly sensitive to the concentration of dsRNAs experienced by the animals on the plates. This high sensitivity may explain the quantitative variability on different days of assays on N2 (e.g. Figure 2 and Table S1).

We note that the number of progeny in the negative controls at 23°C was consistently lower in species of the Drosophilae group compared to those of the Elegans super-group. The number of viable progeny, embryonic delay or arrest varied highly among parents. Lowered fecundity may be partly due to inbreeding depression in these obligate outcrossing species. Dead embryos were also observed in some species of the Elegans super-group, particularly those with a male-female mode of reproduction.

In order to determine the direction and pattern of change in sensitivity to ingested Oti-actin dsRNAs within the Caenorhabditis genus, we made use of a molecular phylogeny of the genus [17]. The pattern of sensitivity and insensitivity is complex when the phylogenetic relationships in the Caenorhabditis genus are taken into account. The state at the base of the tree is unclear. C. sp. 1 was not tested here but was previously shown to be sensitive to external ama-1/pol-2 dsRNAs. A loss of sensitivity in the branch leading to C. plicata is the most parsimonious hypothesis, affecting both the response to ingested Oti-actin and C. plicata ama-1/pol-2 dsRNAs. In the rest of the tree, some pattern emerges. Species in three clades are insensitive: 1) the current sister clade of C. elegans, i.e. eight species including C. briggsae, C. remanei and C. brenneri (except for a weak sensitivity in C. sp. 16); 2) the clade including C. sp. 17, C. sp. 18 and C. sp. 19; 3) the clade of five species including C. angaria and C. drosophilae. From the relationships among sensitive and insensitive species, the most parsimonious possibility is that these three clades have independently lost the ability to respond to ingested Oti-actin dsRNAs. However, gains in the branches leading to the clades 1) C. sp. 20; 2) C. sp. 6 and C. sp. 13; 3) C. sp. 7 and C. sp. 14; 4) C. elegans, and possibly 5) C. sp. 15 depending on its position are also possible, or a combination of gains and losses. In any case, a minimum of four character changes occurred in the Caenorhabditis genus, and of six when including the weak sensitivity of C. sp. 16 and intraspecific evolution in C. elegans.

In C. elegans, different wild isolates were previously shown to be variably sensitive to either germ line or somatic RNAi [9], [30]. The C. elegans N2 reference strain is sensitive in all tissues, albeit to different degrees, e.g. neurons are poorly responsive [31], [32]. By contrast, the C. elegans CB4856 wild isolate is insensitive to germ line RNAi [30]. The C. elegans JU1580 wild isolate was previously found to be insensitive to somatic RNAi (against muscle Cel-unc-22 and GFP; [9], [30]) but sensitive to germ line RNAi (Cel-pos-1). We found that JU1580 is insensitive to Oti-actin dsRNAs (Figure 3 and Table S1) for all tested phenotypes: deformed larvae, dead embryos and sterile mothers (Table S1). This raises the possibility that the scored phenotypes after Oti-actin RNAi are solely the result of somatic inactivation of actin gene expression.

By contrast with the intraspecific variation in C. elegans, we find no variation within C. briggsae, nor with its close relatives: none of the three tested C. briggsae isolates was sensitive to external actin dsRNAs, nor were the most closely related species to C. briggsae, C. spp. 9 and 5 (Figure 3).

Further tests of RNAi sensitivity in a subset of species

Concerning the species that are insensitive to ingested Oti-actin dsRNAs, we further sought to test whether (1) the external administration of dsRNAs or (2) the efficiency of RNAi against the Oti-actin gene (irrespective of its mode of administration) were defective. Previous work had shown that C. briggsae, C. remanei and C. brenneri do not respond to a variety of ingested dsRNAs yet are capable of responding to injected dsRNAs [5]. Among the new species in the same clade, we tested the hermaphroditic C. sp. 11 for its ability to respond to internally administered dsRNAs by injection of Oti-actin dsRNAs into the gonad. We found C. sp. 11 to be sensitive to these internally administered dsRNAs.This rules out a loss of sensitivity due to the evolution of actin gene sequences. It is thus most likely that the whole clade of eight species is defective in the entry of dsRNAs yet has maintained a response to internally administered dsRNAs, allowing for RNA interference studies using injection. In the case of C. brenneri, only injection into the gonad and not into the body cavity was previously found to be effective, suggesting a further inability for systemic spread among tissues in at least in this species [5].

Concerning species of the Drosophilae super-group that are defective in the reponse to external actin dsRNAs, Winston et al. [5] had shown that C. sp. 2 was capable of responding to injected C. sp. 2 ama-1 dsRNAs. We focused on its sister species, C. drosophilae, which is of special interest because of its interesting ecology [33] and on the recently described C. angaria (formerly called C. sp. 3) [34]. Similar to the results in C. sp. 2, we found C. drosophilae to be insensitive to Cdr-ama-1 dsRNAs administered by feeding, but sensitive (although not highly) to the corresponding Cdr-ama-1 dsRNA fragment administered by injection into the gonad. Like C. briggsae, C. drosophilae thus appears to be defective in responding to external but not to internal dsRNAs. Alternatively, a dose effect is possible, assuming that injection delivers a higher dose of dsRNAs than the feeding method.

For C. angaria, we found that external administration of Can-ama-1 dsRNAs by feeding - unlike that of Oti-actin dsRNAs - was highly effective. This result is surprising, because Winston et al. [5] saw no effect when soaking the animals in Can-ama-1 dsRNAs. A dose effect, due to the dsRNA length or the soaking versus feeding mode of administration, could explain this discrepancy. Our own results showing a difference in sensitivity to ingested Oti-actin versus Can-ama-1 dsRNAs may be explained in two ways: i) a better efficiency of, and/or higher dose sensitivity of the inactivation of Can-ama-1 compared to that of Oti-actin; ii) a RNAi competency in only some tissues. Concerning the former hypothesis, upon inspection, the Oti-actin fragment sequence is less similar to the homologous C. angaria sequence (85% identity) than to those of C. elegans (90% identity), which may cause a partial silencing. Consistent with this hypothesis, Oti-actin dsRNAs can trigger a response when injected, yet with low efficiency (Figures 3 and 4). Concerning the latter hypothesis, C. angaria like C. elegans JU1580 ([10]; see above) may be specifically unable to raise a RNAi response in somatic tissues, rather than being sensitive to the mode of dsRNA administration. The inactivation of further genes in C. angaria will be necessary to distinguish between these alternatives. In any case, this species appears sensitive to some ingested dsRNAs.

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Figure 4. RNAi sensitivity of C. sp. 11, C. drosophilae and C. angaria: feeding versus injection.

(A) C. sp. 11 JU1373 was found to be insensitive to Oti-actin RNAi using the feeding protocol (a), but sensitive to Oti-actin dsRNAs introduced by injection (b). (B) C. drosophilae DF5077 was found to be insensitive to RNAi by feeding using both Oti-actin (a) and Cdr-ama-1 (b), but sensitive to Oti-actin dsRNAs introduced by injection (c). C. drosophilae has a smaller brood size than C. elegans in all experiments. (C) C. angaria RGD1 was found to be insensitive to RNAi by feeding using Oti-actin (a) but not Can-ama-1 (b). (c) Compared to control animals injected with Cel-unc-22 dsRNAs, C. angaria RGD1 animals injected with actin dsRNAs showed a significantly reduced number of larvae in their progeny. Injections of actin dsRNAs into C. angaria RGD1 animals were performed in two replicate experiments (shown in blue and red). The proportion of injected P0 that became fully sterile is reported below. Results for C. elegans are shown as positive control for the efficiency of the dsRNA treatment.

https://doi.org/10.1371/journal.pone.0029811.g004

Production of C. briggsae and C. remanei strains that are sensitive to external dsRNAs

Species in the current sister clade to C. elegans were found to be insensitive to ingested dsRNAs. This clade includes species with sequenced genomes where RNAi would be a valuable tool [35] (www.wormbase.org). The RNA interference response could be activated when the dsRNAs were injected [5] (Figure 3). However, RNAi by feeding has several advantages for high-throughput screens [19] and for the screening of late developmental phenotypes by controlling the timing of dsRNA application. We thus set out to render some of the refractory Caenorhabditis species sensitive to dsRNAs administered by feeding, as a tool for further functional studies. This was previously achieved for C. briggsae by transforming a wild-type strain with Cel-SID-2::GFP [5]. The resulting C. briggsae line shows a Roller phenotype due to the rol-6(d)/pRF4 transformation marker and displays intestinal GFP from the Cel-SID-2::GFP transgene. Each of these two features may render some other relevant phenotypes hard to score. In C. briggsae, we thus wished to obtain a non-Roller transgenic line that would help in the scoring of many phenotypes.

As in [5], we could “rescue” C. briggsae insensitivity to ingested dsRNAs with Cel-sid-2, using a simple PCR product of the Cel-sid-2 gene instead of a fusion construct to GFP (see Materials and Methods). We first obtained an extra-chromosomal transgenic line in the C. briggsae AF16 wild background expressing Cel-sid-2 and the Cel-myo-2::DsRed2 marker. After irradiation and backcrosses to C. briggsae AF16, we derived two integrated transgenic strains, JU1018 and JU1076. Both strains were tested for their response to ingested dsRNAs (Table S2). JU1018 was found to be most sensitive to ingested Oti-actin dsRNAs, in fact more sensitive than the reference wild-type C. elegans strain N2; however, in the negative control, JU1018 displayed a somewhat low fertility. JU1076 was more fertile, but somewhat less sensitive to RNAi.

We next tried to similarly render C. remanei sensitive to ingested dsRNAs. We obtained the JU1184 strain by injection of Cel-sid-2 and Cel-myo-2::DsRed2 into the C. remanei PB4641 inbred strain, yielding the mfEx34 transgene. This transgene is transmitted at a 100% frequency and may thus be a spontaneous integrant. This strain displayed a relatively low fertility that rendered the actin test difficult to interpret (since it is based on progeny number) (Table S2). We therefore cloned into the RNAi feeding vector a fragment of the endogenous Cre-unc-22 gene, whose inactivation was predicted to produce a specific Twitcher phenotype [36]. Indeed, whereas we did not observe any Twitcher worms in the control conditions, over 80% of JU1184 progeny raised on bacteria expressing Cre-unc-22 dsRNAs did twitch (Figure 5). We have thus obtained a C. remanei transgenic line that is sensitive to external dsRNAs and may be used for RNA interference experiments.

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Figure 5. Complementing C. briggsae and C. remanei with Cel-sid-2.

(A) Sensitivity to ingested Oti-actin RNAi of two C. briggsae integrated transgenic lines, JU1018 and JU1076, transformed with Cel-sid-2 genomic DNA and a myo-2::DsRed2 marker. Both lines are rendered sensitive to external actin dsRNA application, compared to the reference strain AF16. JU1018 has a lower brood size than the other lines. Statistical comparisons were made to the results of control experiments with Cel-rol-6 or Cbr-lin-12 dsRNA using a Mann-Whitney-Wilcoxon rank sum test on the number of normal larval progeny of each parent. Note that this test is very conservative and has low power. The significance of the difference is depicted as follows: (NS) non-significant, (*) 0.01<p<0.05, (**) 0.001<p<0.01, (***) p<0.001. Error bars indicate the standard error of the mean over individuals (n = 6–12, see Table S1 for details). (B) Sensitivity to Cre-unc-22 RNAi by feeding of a C. remanei transgenic line, JU1184, transformed with Cel-sid-2 genomic DNA and a myo-2::DsRed2 marker. The transgenic strain C. remanei JU1184 displayed the characteristic twitching phenotype when Cre-unc-22 dsRNAs were administered by feeding, whereas no twitcher was seen in the reference strain C. remanei PB4641. χ² test: p = 10⁻⁷⁵.

https://doi.org/10.1371/journal.pone.0029811.g005

Using a similar protocol, we obtained C. sp. 11 transgenic animals carrying the Cel-sid-2 gene. However, we failed to see a strong improvement in external RNAi efficiency when feeding three of these lines with Oti-actin dsRNA expressing bacteria (Table S2). The reasons for this failure of complementing C. sp. 11 with Cel-sid-2 may include insufficient transgene expression or the occurrence of another defective step in the RNAi pathway of this species, for example systemic spread as in C. brenneri [5].

Transgenesis in different Caenorhabditis species

To our knowledge, this is the first report of transgenesis of C. remanei and in the newly isolated selfing C. sp. 11. We further tested whether we could transform Caenorhabditis species using the injection protocol developed for C. elegans, which leads to the formation of extra-chromosomal arrays containing the injected sequences [22]. As transformation markers, we tried a number of C. elegans-based sequences able to induce dominant phenotypes, namely a Roller phenotype or GFP expression (Table 1).

Most tested Caenorhabditis species could be transformed (but see below); however, not all markers were expressed in all species. For example the hsp-16-18::GFP marker was not expressed after heat-shock in C. brenneri nor C. angaria (Table 1A, B, D). By contrast, the sur-5::GFP marker (wide somatic cell expression) or the myo-2::GFP marker (pharyngeal expression) could be successfully used in all tested species of the Elegans and Drosophilae supergroups, including C. brenneri, C. angaria and C. sp. 2. In addition to C. remanei, C. brenneri, C. angaria and C. sp. 2 (Table 1), we could transform C. sp. 11 (Table S2), C. sp. 9 and C. sp. 5 using myo-2::GFP or myo-2::DsRed2 (data not shown). The extra-chromosome could be visualized by Hoechst staining of oocytes in C. angaria (data not shown), showing that in this species of the Drosophilae supergroup as in C. elegans, transgenic lines were obtained by formation of an additional chromosome.

The most basally branching C. sp. 1 could not be transformed using sur-5::GFP nor myo-2::GFP markers. It is unclear whether transgenesis per se or the expression of the markers were unsuccessful. C. sp. 1 displays a smaller body size and has a single functional gonadal arm, which renders injections more difficult. In addition, few progeny was laid after injection. The absence of transformed progeny may be attributed to these complicating factors rather than the intrinsic inability of C. sp. 1 to form extra-chromosomes. In C. plicata, expression of the sur-5::GFP could be observed in the F1 generation after injection and the lack of transformants may simply reflect an insufficient number of injected animals (Table 1).

Concerning the efficiency of transformation, C. elegans, and to a lesser extent C. briggsae, were transformed with a higher frequency of success than other tested species of the Elegans and Drosophilae groups, such as C. remanei, C. brenneri, C. angaria and C. sp. 2 (Table 1). In these latter species, more animals need to be injected to yield the same number of transgenic strains.

Among the transgenic lines, we observed in C. remanei and C. brenneri several lines with 100% transmission of the marker, suggesting direct integration in the genome (Table 1 and the Cel-sid-2-expressing C. remanei strain mentioned above). The proportion of direct integration over extra-chromosome formation appears higher in these species, which is perhaps accounted for by the low efficiency of extra-chromosome formation.

We thus could transform most tested species of the Caenorhabditis genus, using C. elegans markers and protocols. This contrasts with the difficulty in transforming nematodes of other genera, such as Oscheius tipulae [37] or Pristionchus pacificus [38].